The Interactive Fly
Zygotically transcribed genes

DNA Replication enzymes and protein cofactors

How does DNA replication occur?

Chromatin state marks cell-type- and gender-specific replication of the Drosophila genome

Genetic organization of interphase chromosome bands and interbands in Drosophila melanogaster

Histone H4K20 tri-methylation at late-firing origins ensures timely heterochromatin replication

Similarity in replication timing between polytene and diploid cells is associated with the organization of the Drosophila genome

Chromatin conformation and transcriptional activity are permissive regulators of DNA replication initiation in Drosophila

Dynamic changes in ORC localization and replication fork progression during tissue differentiation

Regulatory functions and chromatin loading dynamics of linker histone H1 during endoreplication in Drosophila

Rif1 inhibits replication fork progression and controls DNA copy number in Drosophila

DNA polymerases and subunits

Origin recognition complex

Mini-chromosome maintenance family

DNA replication factor A complex - a single stranded DNA-binding protein complex

Other proteins

anti-silencing factor 1
histone chaperone that assists in chromatin assembly and remodeling during DNA replication, transcription activation, and gene silencing

AAA-superfamily ATP helicase involved in initiation of DNA replication - involved in the formation of the prereplicative complex -
checkpoint protein involved in controlling the G2/M transition

Disc proliferation abnormal
An MCM4 homolog - component of licensing factor

double parked
conserved protein required for DNA replicaton

DNA ligase I
Joins Okazaki fragments

DNA ligase II
Functions in repair (?)

DP transcription factor
transcription factor - obligate dimerization partner of E2f1 and E2f2 - required for normal cell proliferation,
optimal DNA synthesis, and efficient G2/M progression

limits DNA replication by preventing Mcm loading onto chromatin - interacts with Double parked (Drosophila cdt1)

Transcription factor functioning in DNA repair and apoptosis

Proliferating cell nuclear antigen (PCNA) (common alternative name: Mutagen-sensitive 209)
Polymerase-delta/epsilon processivity factor

RNase H1
Involved in Okazaki fragment maturation

Suppressor of Under-Replication
SNF2-domain protein - inhibits replication fork progression to promote DNA underreplication< - binds to H1
which is required for SuUR binding to chromatin in vivo - interacts with Rif1 which has a direct role in copy number control

Topoisomerase 1
Unlinks parental strands - involved in replication

Topoisomerase 2
Unlinks parental strands and progeny duplexes

How does DNA replication take place?

Why should a developmental biologist be interested in DNA replication? There are at least three reasons. (1) For a given origin of replication, there is a link between gene expression and timing of DNA replication, and understanding the basis of this link is important. (2) The mechanism of gene replication is by necessity involved with the restructuring of chromatin and the regulatory implications of that event. (3) There are fail-safe mechanisms to ensure that each origin of replication fires only once per cell cycle, and these mechanisms involve an interaction of cyclins with licensing factor, a chromatin component. Thus, in the future there will be a developing understanding of the relationship between cell cycle, DNA replication, chromatin components, and changes in gene expression.

The first signal for initiation of replication involves replication licensing factor (RLF), which 'licenses' replication origins by putting them into an initiation-competent state. The second signal, S-phase promoting factor, induces licensed origins to initiate, and in doing so removes the license. RLF of Xenopus can be separated into two essential components, RLF-M and RLF-B, both of which are required for licensing. RLF-M, a fraction containing members of the minichromosome maintenance family, associates with chromatin prior to replication but is removed during replication. Drosophila MCM2 and MCM4 homologs have been identified (See disc proliferation abnormal for information about both of these). RLF-M's reassociation with chromatin requires passage through mitosis. RLF-M requires RLF-B, an as yet uncharacterized fraction, for binding RLF-M to DNA. Apparently RLF binds to origins of replication, but the basis for this binding has not yet been characterized (Chong, 1996 and references).

The focus of all replication forks is the helicase, which catalyzes the transition from double- to single-stranded DNA. In eukaryotes, the identification of the enzyme that acts at chromosomal replication forks awaits further investigation, but the SV40 T antigen fulfills the helicase function in SV40 replication. The origin of replication is selected and identified in yeast by the origin recognition complex, of which one Drosophila homolog (Orc2) has been identified. Identification of other components of the ORC and unraveling the nature of DNA sequences at the origin are currently very active subjects of research.

The replication process is semidiscontinuous. Proteins comprising the replication machine act in concert to unwind the parental strands and carry out the simultaneous synthesis of the two progeny strands. Both progeny strands are synthesized in the 5' to 3' direction, but since parental DNA strands are antiparallel, two distinct mechanisms of DNA synthesis are required. One of the two progeny strands (the leading strand) is synthesized continuously in the direction of fork movement. The other (the lagging strand) is synthesized discontinuously in the direction opposite to fork movement. Discontinuous DNA synthesis on the lagging-strand templates involves the related synthesis of oligoribonucleotide primers, which are then elongated into short DNA chains (Okazaki fragments). Following their synthesis, Okazaki fragements are processed to remove the RNA primers and joined together to form an interrupted progeny strand (Brush, 1996).

As the replication fork advances, a helix-destabilizing protein is required to maintain the single-stranded DNA structure that serves as template for RNA priming and DNA synthesis. In eukaryotes, Replication protein A performs this function. This phosphoprotein consists of three subunits.

DNA synthesis is initiated by the bifunctional pol-alpha:primase complex, a heterotetrameric phosphoprotein. The primase activity resides in the smallest subunit and is tightly associated with the next largest, which is thought to tether the primase to the catalytic subunit. The remaining subunit has no known catalytic function, but it may contribute to recruitment of pol-alpha:primase to the replication fork. The main function of pol-alpha:primase is to serve as a priming enzyme. The primase catalyzes the synthesis of complementary oligoribonucleotides, which are then extended a short distance by the polymerase activity. The pol-alpha:primase serves exclusively to initiate DNA synthesis on the lagging strand, and dissociation from the DNA provides a primer terminus for assembly of the PCNA/pol-delta complex, which serves to extend the RNA/DNA primers originally synthesized by pol-alpha:primase.

The heterodimeric DNA polymerase-delta is involved in the elongation stage of DNA replication, acting both on the leading and lagging strands. Unlike pol-alpha:primase, the polymerase-delta does not act alone but requires the action of two auxiliary factors, the multisubunit replication factor C (RF-C) which binds to the primer terminus of the lagging strand immediately after RNA/DNA primer synthesis has been completed by pol-alpha:primase, allowing the subsequent assembly of a functional pol-delta complex. Once bound the primer-template junction, RF-C loads PCNA onto the DNA in an energy-dependent process. PCNA then functions as a processivity factor, acting as a clamp to prevent sliding of polymerase in the reverse direction. RNase H1 acts with another enzyme to remove the RNA primer used to initiate the lagging strand, while DNA ligase I joins the nacent DNA fragments to complete the synthesis of the lagging strand.

It is not clear how pol-alpha:primase synthesized DNA is proofread, as this enzyme has no exonuclease activity. the catalytic subunit of polymerase-delta contains a 3' to 5' exonuclease activity, functioning in proofreading, which allows high-fidelity DNA synthesis (Brush, 1996).

Chromatin state marks cell-type- and gender-specific replication of the Drosophila genome

Duplication of eukaryotic genomes during S phase is coordinated in space and time. In order to identify zones of initiation and cell-type- as well as gender-specific plasticity of DNA replication, replication timing, histone acetylation, and transcription was profiled throughout the Drosophila genome. Two waves of replication initiation were identified with many distinct zones firing in early-S phase, along with multiple, less defined peaks at the end of S phase, suggesting that initiation becomes more promiscuous in late-S phase. A comparison of different cell types revealed widespread plasticity of replication timing on autosomes. Most occur in large regions, but only half coincide with local differences in transcription. In contrast to confined autosomal differences, a global shift in replication timing occurs throughout the single male X chromosome. Unlike in females, the dosage-compensated X chromosome replicates almost exclusively early. This difference occurs at sites that are not transcriptionally hyperactivated, but show increased acetylation of Lys 16 of histone H4 (H4K16ac). This suggests a transcription-independent, yet chromosome-wide process related to chromatin. Importantly, H4K16ac is also enriched at initiation zones as well as early replicating regions on autosomes during S phase. Together, this study reveals novel organizational principles of DNA replication of the Drosophila genome and suggests that H4K16ac is more closely correlated with replication timing than is transcription (Schwaiger, 2009).

The high resolution of these replication timing profiles allowed zones of replication initiation to be identified throughout S phase, that were confirmed in combination with measuring small nascent strand abundance. This revealed that sites of early initiation are rather distinct, which manifests in the timing profile as single peaks or a few peaks clustered together in early-S phase, followed by long stretches of the replication timing profile without changes in slope. Late initiation zones often reside in close proximity to other late initiation zones. The feature of distinct peaks of early initiation in Drosophila is very distinct from mammalian genomes (Hiratani, 2008), where many more sites of initiation of similar timing are clustered together, resulting in large regions up to several megabase pairs of early replication timing (Schwaiger, 2009).

Interestingly, the frequency of initiation appears discontinuous with high rates in early-S, a reduced frequency in mid-S, and again increased appearance of initiation sites in late-S phase. The high frequency and proximity of late-firing initiation zones suggest that late regions are replicated by many proximal late-firing origins of replication. This finding is particularly interesting in light of a recent report that suggested the absence of a checkpoint to control for the completion of DNA replication before mitosis (Torres-Rosell, 2007). This would in turn require a mechanism that mediates rapid replication of unreplicated regions in late-S phase, which could be achieved by a promiscuous activation of many proximal origins. Interestingly, replicative stress that reduces replication fork progression leads to a decrease in inter-origin distance through activation of normally dormant origins. It is conceivable that a similar situation is encountered in late replicating regions (Schwaiger, 2009).

Since the previously reported correlation between replication timing and transcription in Drosophila was not absolute, the percentage of the genome that replicates in a tissue-specific fashion remained to be tested quantitatively. For example, the general correlation could be driven by housekeeping genes that are active in most cells, resulting in a uniform replication timing program. This study showed that dynamic replication timing differs significantly between two Drosophila cell types, affecting at least 20% of autosomal DNA. It was also shown by two different methodologies that this plasticity of DNA replication coincides with transcription differences in only half of all cases (Schwaiger, 2009).

Early replication was shown previously to correlate with transcription levels over 180 kb, leading to the suggestion that replication timing integrates transcription over large regions. Consistent with this model, it was found that dynamic replication timing often occurs in large (~100 kb) regions encompassing many genes. Interestingly, genes with related function often cluster together in the Drosophila genome, and such clusters tend to be similarly 100 kb in size. In mammalian genomes, this clustering appears functionally related to chromatin structure, suggesting that widespread open chromatin at developmentally regulated multigene loci could lead to early replication or vice versa. This, in turn, might increase the potential of gene expression over large regions as in the case of genes important for wing disc development in Cl8 cells, where early replication could render the locus poised for activation (Schwaiger, 2009).

Localized differences in gene expression of a fraction of genes in a large region might also account for replication timing differences. Indeed, some, but not all, genes in differentially replicating regions are strongly differentially expressed between the two cell types. Thus, while gene expression could account for much of the observed changes on autosomes, a considerable fraction does not display transcriptional changes. It seems unlikely that the analysis missed such changes since noncoding transcription was measured as well as RNA polymerase abundance (Schwaiger, 2009).

The relation between replication timing and chromatin structure has been controversial. Transcription itself involves an opening of chromatin structure, and thus early replication could in many situations be downstream from transcriptional activation. However, previous work using injected plasmids suggested a role for early replication in mediating increased levels of histone acetylation. This led to a model in which replication timing mediates an open chromatin structure required for transcription. This suggestion is compatible with the genome-wide analysis, where a preferential location of H4K16ac was observed not only to active genes, but also to early replicating regions that are not transcribed. It is possible that early replication and elevated H4K16ac at inactive genes will result in a more open chromatin confirmation compared with late replicating inactive genes. This might render them more responsive to downstream activating cues, and thus replication timing could modulate the sensitivity to activators. This process could also function in maintenance of an active state through cell division. Importantly, however, this mechanism does not override the parallel process of transcription-coupled acetylation, as late replicating genes that are actively transcribed are still hyperacetylated (Schwaiger, 2009).

Interestingly, a strong abundance of H4K16ac was observed at sites of initiation during S phase. Several single-gene studies have suggested a positive function of histone acteylation for origin activity. Other reports, however, did not support this model. Recent maps of human replication initiation suggest that early origins are marked by H3K9/K14 acetylation (Lucas, 2007). However, no genome-wide correlation between active chromatin marks and early origin firing was observed in S. cerevisiae, where specific sequences function as origins of replication. This study identified a preferential localization of H4K16ac to initiation zones throughout the Drosophila genome compatible with a function of acetylation. In this study, focus was placed on acetylation of H4K16 because this residue has been functionally linked to higher-order chromatin compaction and chromatin opening on the dosage-compensated X in Drosophila (Schwaiger, 2009).

It has been proposed that origins of replication lie frequently between promoters of active genes, which would make transcription and replication fork progression co-oriented. Furthermore, transcription and replication are thought to be coordinated in the nucleus to be spatially and temporally separated. It thus seems plausible that the enrichment of H4K16ac in initiation zones reflects location between highly acetylated, active promoters. According to this model, proximity to active promoters would result in an open chromatin confirmation through increased H4K16ac, which in turn enhances origin firing (Schwaiger, 2009).

Importantly, however, enrichment for H4K16ac was observed at initiation zones that are not proximal to active genes, arguing against a simple process that is solely transcription-coupled. Open chromatin structure, reflected and potentially even mediated by H4K16ac, could make DNA more accessible for efficient initiation of DNA replication and thus provide a sequence-independent component that could contribute to origin localization and activity. While these are testable models, they do require a fine-mapping of actual origins at a resolution higher than the current detection of zones of initiation at the level of several kilobases (Schwaiger, 2009).

This analysis reveals the almost complete absence of late replication on the single X chromosome in male Drosophila cells. About 90% of female late replicating regions on the X replicate early in males, while autosomes show no advanced replication. Such chromosome-wide advance in replication timing has not been observed previously. In mammals, transcriptional inactivation of one of the female X chromosomes correlates with its late replication, reflecting the efficient silencing of this chromosome and increased chromatin compaction. In contrast, dosage compensation in flies involves the twofold up-regulation of genes already active in females and an open chromatin state mediated by H4K16ac. Interestingly, this study showed that advanced replication of the dosage-compensated X occurs mostly outside of transcriptionally activated regions and thus is unlikely to be accounted for by transcriptional changes. Importantly, the local increases in H4K16ac, which are detected throughout the male X chromosome, can be directly related to this loss of late replication. Reduction of the responsible Histone-Acetyltransferase Mof leads to a block in cell division, making it difficult to test this model. Notably, slightly delayed replication of the X chromosome was detected in the few cells that were in S phase in the knockdown population. While this is compatible with a model that Mof-mediated H4K16 acetylation advances replication of intergenic regions on the male X chromosome, the predominant effect on the cell cycle precluded further analysis (Schwaiger, 2009).

This suggests a transcription-independent, chromatin-dependent process, which leads to early replication chromosome-wide. While this likely reflects a different chromatin compaction, it is tempting to speculate that it also reflects a particular nuclear organization as the dosage-compensated X chromosome has been shown to associate directly with nuclear pores (Schwaiger, 2009).

Together these findings provide new principles of the replication timing program of the Drosophila genome and its dynamics relative to histone acetylation and transcription. The data further support a model in which open chromatin structure is a general feature of early replication timing and could potentially even advance replication of entire chromosomes (Schwaiger, 2009).

Genetic organization of interphase chromosome bands and interbands in Drosophila melanogaster

Drosophila melanogaster polytene chromosomes display specific banding pattern; the underlying genetic organization of this pattern has remained elusive for many years. This paper analyzed 32 cytology-mapped polytene chromosome interbands. Molecular locations of these interbands was estimated, their molecular and genetic organization was described and it was demonstrated that polytene chromosome interbands contain the 5' ends of housekeeping genes. As a rule, interbands display preferential 'head-to-head' orientation of genes. They are enriched for 'broad' class promoters characteristic of housekeeping genes and associate with open chromatin proteins and Origin Recognition Complex (ORC) components. In two regions, 10A and 100B, coding sequences of genes whose 5'-ends reside in interbands map to constantly loosely compacted, early-replicating, so-called 'grey' bands. Comparison of expression patterns of genes mapping to late-replicating dense bands vs genes whose promoter regions map to interbands shows that the former are generally tissue-specific, whereas the latter are represented by ubiquitously active genes. Analysis of RNA-seq data (modENCODE-FlyBase) indicates that transcripts from interband-mapping genes are present in most tissues and cell lines studied, across most developmental stages and upon various treatment conditions. A special algorithm was developed to computationally process protein localization data generated by the modENCODE project; it was shown that Drosophila genome has about 5700 sites that demonstrate all the features shared by the interbands cytologically mapped to date (Zhimulev, 2014. PubMed ID: 25072930).

DNA copy-number control through inhibition of replication fork progression

Proper control of DNA replication is essential to ensure faithful transmission of genetic material and prevent chromosomal aberrations that can drive cancer progression and developmental disorders. DNA replication is regulated primarily at the level of initiation and is under strict cell-cycle regulation. Importantly, DNA replication is highly influenced by developmental cues. In Drosophila, specific regions of the genome are repressed for DNA replication during differentiation by the SNF2 domain-containing protein Suppressor of Under-Replication (SuUR) through an unknown mechanism. This study demonstrates that SuUR is recruited to active replication forks and mediates the repression of DNA replication by directly inhibiting replication fork progression instead of functioning as a replication fork barrier. Mass spectrometry identification of SUUR-associated proteins identified the replicative helicase member CDC45 as a SUUR-associated protein, supporting a role for SUUR directly at replication forks. These results reveal that control of eukaryotic DNA copy number can occur through the inhibition of replication fork progression (Nordman, 2014: PubMed).

Histone H4K20 tri-methylation at late-firing origins ensures timely heterochromatin replication

Among other targets, the protein lysine methyltransferase PR-Set7 (see Drosophila SET domain containing 7) induces histone H4 lysine 20 monomethylation (H4K20me1), which is the substrate for further methylation by the Suv4-20h methyltransferase. Although these enzymes have been implicated in control of replication origins, the specific contribution of H4K20 methylation to DNA replication remains unclear. This study shows that H4K20 mutation in mammalian cells, unlike in Drosophila, partially impairs S-phase progression and protects from DNA re-replication induced by stabilization of PR-Set7. Using Epstein-Barr virus-derived episomes, it was further demonstrated that conversion of H4K20me1 to higher H4K20me2/3 states by Suv4-20h is not sufficient to define an efficient origin per se, but rather serves as an enhancer for MCM2-7 helicase (see Drosophila MCM5) loading and replication activation at defined origins. Consistent with this, it was found that Suv4-20h-mediated H4K20 tri-methylation (H4K20me3) is required to sustain the licensing and activity of a subset of ORCA/LRWD1-associated origins, which ensure proper replication timing of late-replicating heterochromatin domains. Altogether, these results reveal Suv4-20h-mediated H4K20 tri-methylation as a critical determinant in the selection of active replication initiation sites in heterochromatin regions of mammalian genomes (Brustel, 2017).

Similarity in replication timing between polytene and diploid cells is associated with the organization of the Drosophila genome

Morphologically, polytene chromosomes of Drosophila melanogaster consist of compact 'black' bands (ruby bands, rb-bands) alternating with less compact 'grey' bands and interbands. This study developed a comprehensive approach that combines cytological mapping data of FlyBase-annotated genes and novel tools for predicting cytogenetic features of chromosomes on the basis of their protein composition and determined the genomic coordinates for all black bands of polytene chromosome 2R. By a PCNA immunostaining assay, the replication timetable was obtained for all the bands mapped. The results allowed comparison of replication timing between polytene chromosomes in salivary glands and chromosomes from cultured diploid cell lines and to observe a substantial similarity in the global replication patterns at the band resolution level. In both kinds of chromosomes, the intervals between black bands correspond to early replication initiation zones. Black bands are depleted of replication initiation events and are characterized by a gradient of replication timing; therefore, the time of replication completion correlates with the band length. The bands are characterized by low gene density, contain predominantly tissue-specific genes, and are represented by silent chromatin types in various tissues. The borders of black bands correspond well to the borders of topological domains as well as to the borders of the zones showing H3K27me3, SUUR, and LAMIN enrichment. In conclusion, the characteristic pattern of polytene chromosomes reflects partitioning of the Drosophila genome into two global types of domains with contrasting properties. This partitioning is conserved in different tissues and determines replication timing in Drosophila (Kolesnikova, 2018).

This work used the four-state chromatin model, previously published data on the chromatin localization of proteins, and in situ hybridization of annotated genes and identified the locations of all black bands of polytene chromosome 2R on a genome map (Kolesnikova, 2018).

The special feature of the four-state chromatin model is the generalization of data obtained from four cell lines. This generalization resulted in identification of two chromatin types -- aquamarine and ruby -- which show stable properties in all these cell lines. Starting the current mapping effort with identification of domains that have ruby chromatin in them and that are flanked by aquamarine chromatin, the genome is roughly divided into constitutively active and constitutively inactive zones (Kolesnikova, 2018).

To take into account the tissue-specific features of chromatin organization in salivary gland polytene chromosomes, the morphology of polytene chromosomes, an extensive pool of 'experimental cytology' data, and data on gene expression in salivary glands were examined and, whenever deemed necessary, corrections were introduced into initial predictions. The results of this study revealed that this approach works well (Kolesnikova, 2018).

Yet another unique feature of the four-state chromatin model is that it reveals the chromatin type (specifically, aquamarine) that is enriched with interband-specific proteins; no other models of clustering chromatin proteins can do this. According to the most recent high-resolution Hi-C data from embryos, TAD boundaries correspond with high resolution to polytene chromosome interbands, whereas black and gray bands are the visualization of topological domains with different types of DNA folding. Thus, the four-state chromatin model allows the boundaries of physical (not epigenetic) domains to be found. The work by Hou (2012) clearly indicates that these boundaries are not always the same (Kolesnikova, 2018).

The choice of aquamarine chromatin as potential band boundaries is supported well by comparing coordinates with the distribution of SUUR and H3K27me3 on polytene chromosomes: these two are markers of black bands and display sharp changes on their boundaries. Thus, the approach used in the current work can be conveniently used to identify the coordinates of specific polytene-chromosome black bands with high accuracy. For most black bands, accuracy of 2-10 kb is attainable (Kolesnikova, 2018).

With the band coordinates inferred, a detailed comparison was performed of replication profiles from diploid cells and replication patterns observed in polytene chromosomes and analyzed the properties of black bands (Kolesnikova, 2018).

A considerable degree of similarity in replication timing has been demonstrated between salivary gland polytene chromosomes and diploid cells. In both object types, the zones between black bands correspond to early replication initiation zones. This result is consistent with the observation that most ORC2-binding sites are in aquamarine chromatin corresponding to interbands. Ruby-containing polytene chromosome bands (Rb-bands) in different cell types have a U-shaped replication profile, which implies that replication in them proceeds from the boundaries to the center, leading to a local delay in replication completion, this delay being proportional to the band length (Kolesnikova, 2018).

The averaged boundary replication profiles in INTs that were built from previously published cell culture data are consistent with the prediction that very few sites within the intervals between rb-bands initiate replication in each replication cycle. The typical size of replicons, 80 kb, originating from the early replication initiation zone (data from cell cultures) fits this model well too. Analysis of stretched DNA fibers in D. nasuta polytene chromosomes has revealed that the replicons initiated in the early S phase are each 64 μm in size on average, which should amount to more than 120 kb. The fact that the replicons are that long provides further support to the hypothesis that the replication origins fired during one cycle in a particular DNA molecule should be well spaced. While analyzing replication in partially denatured polytene chromosomes, researchers observed temporal and spatial asynchrony in replication initiation in parallel fibers and proposed that this asynchrony is one of the main reasons for continuous labeling in polytene chromosomes. Although these data come from a Drosophila species irrelevant to this study and the typical sizes and genomic distances may be different to some extent, it is proposed that the organization of replication in polytene chromosomes is conserved across Drosophila species (Kolesnikova, 2018).

Replication patterns change in polytene chromosomes, and these patterns are linked to events in DNA sequences. At the beginning of the S phase, replication is initiated in INTs, which may contain a large number of potential replication initiation sites. In each DNA strand, an initiation event occurs only once per INT in a random interband. Initiation events in different interbands can occur in asynchrony, either in different INTs or in the same INT on the parallel DNA strands of a polytene chromosome. Replication forks move through INTs in the opposite directions from the site of replication initiation and eventually enter the nearest rb-bands. After all INTs have completed replication, the replication fork should be detectable only in rb-bands. This situation is consistent with the observed inverted PCNA pattern, when all black bands produce the signal that the intervals do not (Kolesnikova, 2018).

By analysis of stretched DNA fibers in D. nasuta polytene chromosomes, it has been demonstrated that the rates of replication fork movement in polytene chromosomes during the late S phase are on average one-tenth of those in the early S phase. The authors believe that upon entering polytene chromosome rb-bands, replication forks slow down. That is why, although some rb-bands are shorter than the flanking intervals, all 'black' bands undergoing replication at once. Replication in these bands goes on until the forks moving toward each other meet. In the longest rb-bands, forks fail to meet before the end of the S phase, leading to under-replication. In D. melanogaster, replication rates depend on SUUR. This study demonstrated that all rb-bands are enriched with SUUR both in salivary gland polytene chromosomes and in diploid cells. According to another study, local artificial tethering of SUUR to an early replicating region of a salivary gland polytene chromosome causes delayed replication there. It can be proposed that this protein plays an important role in delayed replication associated with all rb-bands genome-wide (Kolesnikova, 2018).

Evidence exists that the S phase of the endocycle is quite different from that in diploid cells. The former is distinguished by under-replication of a large part of the genome and low expression of genes involved in replication. The presence of the intra-S checkpoint in salivary gland cells is questionable, and so is activation of any late-firing origins. Nevertheless, the results reveal a substantial similarity in replication timing for the euchromatic arm of the whole chromosome. What underlies this similarity is thought to be the organization of the Drosophila genome. The genome consists of alternations of domains capable of initiating early replication (INTs; the interval between black/ruby bands) and domains with the potential to initiate replication late in the S phase. These late domains vary in size, but seldom are they longer than a few hundred kilobases. In diploid cells, these relatively short domains are replicated by replication forks coming from border origins of replication and complete replication before the classic late S phase, which is when late-firing origins activate. Thus, replication of a large portion of a euchromatic arm is, in the classic sense, early replication. Only the most extended bands and regions of pericentric heterochromatin initiate replication in the late S phase. The question of whether replication initiation events occur in the bands is not easy to answer. Schwaiger (2009) analyzed replication profiles and concluded that extended late-replication zones in cell cultures contain origins initiating replication shortly before the end of the S phase. It can be assumed that the replication origins located in rb-bands do not bind all proteins required for independent initiation of replication, and replication on these origins cannot be initiated before a fork comes from outside; these properties are typical of regions showing a U-shaped replication profile in mammals. The same is suggested by recent studies of the genome-wide distribution of the Mcm2-7 helicase complex in D. melanogaster (Kolesnikova, 2018).

Multiple published comparisons of replication profiles for different tissues of the same organism suggest that each cell type has its own schedule of origin activation. One study on individual IH regions indicates that all the 60 analyzed regions are late replicating in cultured cells, but inside those regions, there are local zones of early replication. After artificial induction of transgene expression in IH, there are also local changes in replication timing (Kolesnikova, 2018).

In cell cultures, early-replication zones within rb-bands can be identified by analysis of the outliers in the boxplots of the averaged boundary replication profile. Among all the rb-bands, only two had early-replication zones spanning them from end to end. It can be theorized that in different tissues, most bands similar in size undergo replication within a similar time interval in the S phase, and gene activation in these bands makes the corresponding fragment of the band earlier replicating (Kolesnikova, 2018).

It can be concluded that the alternation of rb-bands and INTs forms the basis of the pattern of replication timing in D. melanogaster. This organization is conserved in eukaryotes. It has been demonstrated that a substantial portion of the mammalian genome represents the alternation of replication initiation zones, in which early master origins lie, and U-shaped replication zones, in which initiation occurs at virtually random positions and in a cascadelike manner, shaping the profile accordingly. The initiation zones are notable for active transcription and high gene density. The boundaries of these zones correspond to those of topological domains (Kolesnikova, 2018).

IH regions represent a separate fraction of black bands, grossly corresponding to the most extended and late-replicating bands (group LR5). This study demonstrated that all rb-bands, including small ones, share a large number of properties with IH regions. This is direct evidence that among all genomic regions, IH regions do not stand out as some special type of sequences. Genes in any rb-band tend to be expressed in a limited number of tissues and, according to GO analysis, these regions are enriched with tissue-specific genes. By contrast, the intervals between black bands are enriched with genes that are highly expressed in most tissues chosen for analysis here. Each rb-band appears as a combination of repressed chromatin types; however, open chromatin can be found in its boundary regions, pointing to a similarity between bands and TADs. It is confirmed that the boundaries of black bands correspond to those of topologically associating domain or sub-domains. TADs represent a stable level of genome organization during development both in mammals and in Drosophila. It has been demonstrated that the partitioning of genomes into physical domains correlates with gene density and transcription distribution. These features are closely associated with replication timing. That late replication correlates with LADs has been demonstrated in both Drosophila and mammals (Kolesnikova, 2018).

The results of this work suggest that Drosophila polytene chromosomes can serve as vivid visualization of the organization of the eukaryotic genome, which is conserved between Drosophila and mammals. The characteristic pattern of polytene chromosomes -- the compacted black bands alternating with less compact grey bands and interbands -- reflects the partitioning of the Drosophila genome into domains with contrasting properties (Kolesnikova, 2018).

Chromatin conformation and transcriptional activity are permissive regulators of DNA replication initiation in Drosophila

Chromatin structure has emerged as a key contributor to spatial and temporal control over the initiation of DNA replication. Nevertheless, a causal relationship between chromatin structure and replication initiation remains elusive. This study combined histone gene engineering and whole-genome sequencing in Drosophila to determine how perturbing chromatin structure affects replication initiation. Most pericentric heterochromatin was found to remain late replicating in H3K9R mutants, even though H3K9R pericentric heterochromatin is depleted of HP1a, more accessible, and transcriptionally active. These data indicate that HP1a loss, increased chromatin accessibility, and elevated transcription do not result in early replication of heterochromatin. Nevertheless, a small amount of pericentric heterochromatin with increased accessibility replicates earlier in H3K9R mutants. Transcription is de-repressed in these regions of advanced replication, but not in those regions of the H3K9R mutant genome that replicate later, suggesting that transcriptional repression may contribute to late replication. This study also explored relationships among chromatin, transcription, and replication in euchromatin by analyzing H4K16R mutants. In Drosophila, the X Chromosome is upregulated 2-fold and replicates earlier in XY males than it does in XX females. This study found that H4K16R mutation prevents normal male development and abrogates hyper-expression and earlier replication of the male X, consistent with previously established genome-wide correlations between transcription and early replication. By contrast, H4K16R females are viable and fertile, indicating that H4K16 modification is dispensable for genome replication and gene expression (Armstrong, 2018).

Dynamic changes in ORC localization and replication fork progression during tissue differentiation

Genomic regions repressed for DNA replication, resulting in either delayed replication in S phase or underreplication in polyploid cells, are thought to be controlled by inhibition of replication origin activation. Studies in Drosophila polytene cells, however, raised the possibility that impeding replication fork progression also plays a major role. This study exploited genomic regions underreplicated (URs) with tissue specificity in Drosophila polytene cells to analyze mechanisms of replication repression. By localizing the Origin Recognition Complex (ORC) in the genome of the larval fat body and comparing this to ORC binding in the salivary gland, sites of ORC binding were found to show extensive tissue specificity. In contrast, there are common domains nearly devoid of ORC in the salivary gland and fat body that also have reduced density of ORC binding sites in diploid cells. Strikingly, domains lacking ORC can still be replicated in some polytene tissues, showing absence of ORC and origins is insufficient to repress replication. Analysis of the width and location of the URs with respect to ORC position indicates that whether or not a genomic region lacking ORC is replicated is controlled by whether replication forks formed outside the region are inhibited. These studies demonstrate that inhibition of replication fork progression can block replication across genomic regions that constitutively lack ORC. Replication fork progression can be inhibited in both tissue-specific and genome region-specific ways. Consequently, when evaluating sources of genome instability it is important to consider altered control of replication forks in response to differentiation (Hue, 2018).

Regulatory functions and chromatin loading dynamics of linker histone H1 during endoreplication in Drosophila

Eukaryotic DNA replicates asynchronously, with discrete genomic loci replicating during different stages of S phase. Drosophila larval tissues undergo endoreplication without cell division, and the latest replicating regions occasionally fail to complete endoreplication, resulting in underreplicated domains of polytene chromosomes. This study shows that linker histone H1 is required for the underreplication (UR) phenomenon in Drosophila salivary glands. H1 directly interacts with the Suppressor of UR (SUUR) protein and is required for SUUR binding to chromatin in vivo. These observations implicate H1 as a critical factor in the formation of underreplicated regions and an upstream effector of SUUR. It was also demonstrated that the localization of H1 in chromatin changes profoundly during the endocycle. At the onset of endocycle S (endo-S) phase, H1 is heavily and specifically loaded into late replicating genomic regions and is then redistributed during the course of endoreplication. The data suggest that cell cycle-dependent chromosome occupancy of H1 is governed by several independent processes. In addition to the ubiquitous replication-related disassembly and reassembly of chromatin, H1 is deposited into chromatin through a novel pathway that is replication-independent, rapid, and locus-specific. This cell cycle-directed dynamic localization of H1 in chromatin may play an important role in the regulation of DNA replication timing (Andreyeva, 2017).

This study demonstrated that virtually all major sites of UR throughout the Drosophila genome exhibit a substantial increase in salivary gland DNA copy number upon depletion of the linker histone H1, thus implicating H1 in the regulation of endoreplication. In control knockdown salivary glands, 46 underreplicated domains were identified. While these regions are in general agreement with previous efforts to map underreplicated domains by less sensitive microarray analyses, fewer underreplicated sites were identified than a recent report that used high-throughput sequencing of salivary gland DNA (Yarosh, 2014). Notably, the underreplicated domains that the current analyses failed to detect represent sites with the weakest degree of UR. One possible source of variation is the distinct technical approach that was used compared with Yarosh (2014), as simultaneous sequencing of a nonpolytenized (embryonic) genome as a means to normalize the reads from underrepresented sequences in polytenized tissues (Yarosh, 2014) likely provides additional sensitivity. Another potential explanation could lie in the relative sequencing depth of the respective assays (approximately fourfold lower in the current study), considered crucial for the analyses of next-generation sequencing data. However, this explanation is less likely, as subsampling of the current reads to much lower depths yielded no appreciable difference in the number and location of identified underreplicated sites or the change in copy number upon H1 knockdown (Andreyeva, 2017).

On average, a moderate knockdown of H1 led to an ~50% copy number gain at the center of underreplicated domains in intercalary heterochromatin (IH; large dense bands scattered in euchromatin comprising clusters of repressed genes. The copy number is not restored to the same degree as that in a SuUR genetic mutant. The difference is likely attributable to the incomplete depletion of H1. In fact, in an independent biological validation experiment that resulted in an ~95% depletion of H1, an almost complete restoration of copy number was observed. The observation of an almost complete reversal of UR in cells depleted of H1 (but still wild type for SuUR) strongly suggests an epistatic mechanism of action in which both H1 and SUUR act together in the same biochemical pathway (Andreyeva, 2017).

This study found that H1 and SUUR are also involved in UR of PH. For instance, both the mapped pericentric regions and TE sequences, which are highly abundant in pericentric regions, exhibit an increase of DNA copy number upon H1 knockdown. The SuURES mutation also results in a robust loss of UR at PH, as measured by changes in DNA copy number at TEs. The abrogation of H1 expression gives rise to a somewhat weaker effect on the UR of PH than that of IH, which is consistent with an almost complete elimination of SUUR protein from polytene chromosome arms in salivary glands depleted of H1 by RNAi but the persistence of residual SUUR at their PH. The role of H1 in maintaining the underreplicated state of PH may be relevant to its important regulatory functions in constitutive heterochromatin, where it recruits Su(var)3-9, facilitates H3K9 methylation, and maintains TEs in a transcriptionally repressed state. Recently, it was proposed that TE repression in ovarian somatic cells involves an H3K9 methylation-independent process through recruitment of H1 by Piwi-piRNA complexes, resulting in reduced chromatin accessibility. The current results also implicate UR of TE sequences in polytenized cells as yet another putative mechanism that contributes to regulation of their expression. Interestingly, it was shown previously that double mutants encompassing both the Su(var)3-9 and SuUR mutant alleles exhibit a synthetically increased predominance of novel band-interband structures at PH compared with the mutation of SuUR alone. While the evidence suggests a relationship between UR and transcriptionally repressive epigenetic states, such as H3K9 methylation, the nature of this relationship remains largely speculative (Andreyeva, 2017).

This study demonstrated that SUUR protein physically interacts with H1 in both a complex mixture of whole-cell extracts that contain endogenous native H1 and recombinant purified H1 polypeptides. Furthermore, the particular structural domains of the two proteins were delimited that are required for the interaction. SUUR protein contains several sequence features that have been implicated in regulation of UR and binding to specific proteins. Although SUUR possesses a putative bromodomain, it contains no identifiable DNA-binding domain, so the mechanism that allows SUUR to exhibit a preference for specific genomic underreplicated loci is unknown. The positively charged central region is both necessary and sufficient to interact with heterochromatin protein 1a (HP1a), which suggests a possible involvement of HP1a in tethering SUUR to H3K9me2/3-rich PH. However, the specific localization of SUUR to underreplicated IH, which is not enriched for H3K9me2/3, remains enigmatic. This study now demonstrates that the central region of SUUR is also sufficient for binding directly to H1 in vitro. Considering that the central region of SUUR is essential for the faithful localization of the protein to chromatin in vivo, including underreplicated IH, it seems likely that H1 directly mediates the tethering of SUUR to chromatin in underreplicated regions (Andreyeva, 2017).

The tripartite structure of H1 provides multiple binding interfaces for interacting proteins and thus allows H1 to mediate several biochemically separable functions in vivo. For instance, the globular domain and proximal 25% of the CTD are required for H1 loading into chromatin, while the proximal 75% of the CTD is needed for normal polytene morphology, H3K9 methylation, and physical interactions with Su(var)3-9. This study discovered a previously unknown function for the distal 25% of the H1 CTD, which is shown to be essential for binding to SUUR. Deletion of this region of H1 results in a near-complete loss of the interaction with SUUR. Thus, in addition to its critical functions in heterochromatin structure and activity, the CTD of H1 is likely also important in facilitating UR (Andreyeva, 2017).

One of the most striking findings in this study is the observation that the genomic occupancy of H1 undergoes profound changes during the endoreplication cycle. It also remains largely mutually exclusive with that of DNA polymerase clamp loader PCNA, which is consistent with the observed depletion of H1 in nascent chromatin compared with mature chromatin (Andreyeva, 2017).

H1 is heavily loaded into late replicating loci at the onset of replication (when these loci are silent for replication). Combined, the current observations indicate that the chromosome distribution of H1 during the endocycle is governed by at least three independent processes. Two of them [replication-dependent (RD) eviction of H1 and RD deposition of H1 after the passage of replication fork] are related to the well-recognized obligatory processes of chromatin disassembly and reassembly during replication. The third pathway, which directs early deposition of H1 into late replicating loci, has not been described previously. This process is (1) replication-independent (RI); (2) locus-specific, with a strong preference for late replicating sites; and (3) apparently more rapid than the RD deposition of H1, since very high levels of H1 occupancy are observed in all nuclei immediately after the initiation of endo-S. It is possible that the RI pathway of H1 loading into chromatin is mediated by a selective recruitment of H1 based on epigenetic core histone modification-dependent mechanisms. For instance, mammalian H1.2 was reported to recognize H3K27me3, and this modification is very abundant in IH (Sher et al. 2012) (Andreyeva, 2017).

Also, the RI mechanism for deposition of H1 probably does not involve de novo nucleosome assembly, as H1 is known to exhibit a mutually exclusive distribution with RI core histone variants, and there is no known nuclear process during early S phase that requires core histone turnover. In the future, it will be interesting to further confirm that RI nucleosome assembly does not take place during early replication in salivary gland polytene chromosomes. Finally, the locus-specific RI deposition of H1 in early endo-S chromatin may be conserved in the normal S phase of diploid tissues, and it will require independent experimentation with sorted mitotically dividing cells to confirm this possibility (Andreyeva, 2017).

This study also provides cytological evidence that the functions of H1 and SUUR are biochemically linked. Specifically, it was demonstrated that SUUR localizes to a subset of H1-positive bands and requires H1 for its precise distribution in polytene chromosomes, nuclear localization, and stability in salivary gland cells. These observations implicate H1 as an upstream effector of SUUR functions in vivo and an essential component of the biological pathway that maintains loci of reduced ploidy in polytenized cells. Importantly, this finding adds to a growing list of biochemical partners of H1 that mediate their chromatin-directed functions in an H1-dependent fashion (Andreyeva, 2017).

Interestingly, even a moderate depletion of H1 (to ~30% of normal) results in a complete removal of SUUR from chromosome arms. Thus, H1-dependent localization of SUUR requires high concentrations of the linker histone in chromatin. This conclusion is also consistent with SUUR colocalization with polytene loci that are the most strongly stained for H1. In contrast, elimination of the H3K9me2 mark from polytene spreads requires very extensive depletion of H1, whereas the moderate depletion of H1 does not strongly affect H3K9 dimethylation in the chromocenter or polytene arms. Therefore, the robust effect of even moderate H1 depletion on SUUR localization in chromatin is unlikely to be mediated indirectly through disorganization of heterochromatin structure (Andreyeva, 2017).

Unexpectedly, the cell cycle-dependent temporal pattern of H1 localization is not identical to that of SUUR. In contrast to H1, SUUR protein (1) is only weakly present in IH during early endo-S phase, (2) achieves the maximal occupancy at IH loci only in the late endo-S, and (3) colocalizes with PCNA at certain sites. The observations made in this study and in previous works can be summarized in the following model for H1-mediated regulation of SUUR association with chromatin. The initiation of the deposition of SUUR in chromosomes is strongly dependent on H1. More specifically, SUUR is preferentially localized to chromatin domains that are highly enriched for H1. For instance, the tremendously elevated concentration of H1 in IH of early endo-S cells promotes and nucleates the initiation of deposition of SUUR into these regions. However, the pattern of SUUR occupancy at these sites does not occur temporally in parallel with that of H1. Initially, the exceptionally high abundance of H1 in late replicating loci during early endo-S is not paralleled by a simultaneous comparable increase of SUUR occupancy. Rather, loading of SUUR into these sites lags significantly behind H1 occupancy. Thus, the rate of SUUR localization to H1-rich IH appears to be much slower than that of the RI deposition of H1 into these loci. After the initial recruitment, further loading of SUUR does not require H1, and SUUR continues (in a slower fashion) to accumulate at IH throughout the endo-S phase even when H1-enriched domains dissipate in the course of DNA endoreplication. The additional loading of SUUR in chromatin is likely facilitated by its self-association through dimerization of the N terminus and physical interactions with the replication fork, as proposed previously. In this fashion, SUUR achieves its maximal concentration in IH loci by the late endo-S (Andreyeva, 2017).

This study has demonstrated that H1 has a pivotal function in the establishment of UR of specific IH loci in polytenized salivary gland cells. The findings that H1 interacts directly with SUUR in vitro and is required for SUUR localization to late replicating IH in polytene chromosomes in vivo strongly suggest that the H1-mediated recruitment of SUUR promotes UR by obstructing replication fork progression in its cognate underreplicated loci but does not affect replication origin firing. However, the remarkable temporal pattern of H1 distribution in endoreplicating polytene chromosomes suggests that it may also play a direct SUUR-independent role in regulation of endoreplication. This is especially plausible considering that the temporal distribution patterns of SUUR and H1 are dissimilar (Andreyeva, 2017).

In contrast to the role of SUUR in slowing down the replication fork progression during late endo-S phase, H1 (acting in the absence of SUUR during early endo-S) may function to repress the initiation of endoreplication, as proposed in several studies. DNA-seq analyses also suggest this mechanism. Compared with the relatively smooth, flat profiles of DNA copy numbers in SuURES mutant salivary glands, the profiles in H1-depleted cells exhibit a jagged, uneven appearance, indicative of aberrant local initiation of replication. Unfortunately, the experimental system (cytological analyzes of salivary glands) cannot be used to further confirm this idea. First, an extensive depletion of H1 results in the loss of polytene morphology; second, since the staging of endo-S progression is based on PCNA staining, a spurious activation of ectopic replication origins would result in an incorrect calling of the stage. To further complicate these analyses, polytenized cells are not amenable to other methods of cell cycle staging, such as fluorescence-activated cell sorting (FACS). In the future, it will be important to examine the role of H1 in regulation of DNA replication timing in sorted Drosophila diploid cells (Andreyeva, 2017).

Rif1 inhibits replication fork progression and controls DNA copy number in Drosophila

Control of DNA copy number is essential to maintain genome stability and ensure proper cell and tissue function. In Drosophila polyploid cells, the SNF2-domain-containing SUUR protein inhibits replication fork progression within specific regions of the genome to promote DNA underreplication. While dissecting the function of SUUR's SNF2 domain, an interaction between SUUR and Rif1 was identified. Rif1 has many roles in DNA metabolism and regulates the replication timing program. Repression of DNA replication is dependent on Rif1. Rif1 localizes to active replication forks in a partially SUUR-dependent manner and directly regulates replication fork progression. Importantly, SUUR associates with replication forks in the absence of Rif1, indicating that Rif1 acts downstream of SUUR to inhibit fork progression. These findings uncover an unrecognized function of the Rif1 protein as a regulator of replication fork progression (Munden, 2018).

The SUUR protein is responsible for promoting underreplication of heterochromatin and many euchromatin regions of the genome. Although SUUR was recently shown to promote underreplication through inhibition of replication fork progression, the underlying molecular mechanism has remained unclear. Through biochemical, genetic, genomic and cytological approaches, this study has found that SUUR recruits Rif1 to replication forks and that Rif1 is responsible for underreplication. This model is supported by several independent lines of evidence. First, SUUR associates with Rif1, and SUUR and Rif1 co-localize at sites of replication. Second, underreplication is dependent on Rif1, although Rif1 mutants have a clear pattern of late replication in endo cycling cells. Third, SUUR localizes to replication forks and heterochromatin in a Rif1 mutant, however, it is unable to inhibit replication fork progression in the absence of Rif1. Fourth, Rif1 controls replication fork progression and phenocopies the effect loss of SUUR function has on replication fork progression. Fifth, SUUR is required for Rif1 localization to replication forks. Critically, using the gene amplification model to separate initiation and elongation of replication, it was shown that Rif1 can affect fork progression without altering the extent of initiation. Based on these observations, this study defines a new function of Rif1 as a regulator of replication fork progression (Munden, 2018).

This work suggests that the SNF2 domain of SUUR is critical for its ability to localize to replication forks. This is based on the observation that deletion of this domain results in a protein that is unable to localize to replication forks, but still localizes to heterochromatin. SUUR has previously been shown to dynamically localize to replication forks during S phase, but constitutively binds to heterochromatin (Kolesnikova, 2013; Nordman, 2014). SUUR associates with HP1 and this interaction occurs between the central region of SUUR and HP1 (Pindyurin, 2008). Therefore, it is speculated that the interaction between SUUR and HP1 is responsible for constitutive SUUR localization to heterochromatin, while a different interaction between the SNF2 domain and a yet to be defined component of the replisome, or replication fork structure itself, recruits SUUR to active replication forks during S phase (Munden, 2018).

Uncoupling of SUUR's ability to associate with replication forks and heterochromatin also provides a new level of mechanistic understanding of underreplication. Overexpression of the C-terminal two-thirds of SUUR is capable of inducing ectopic sites of underreplication. In contrast, overexpression of the SUUR's SNF2 domain, in the presence of endogenous SUUR, suppresses SUUR-mediated underreplication (Kolesnikova, 2005). Together with the data presented in this study, it is suggested that overexpression of the SNF2 domain interferes with recruitment of full-length SUUR to replication forks, by saturating potential SUUR binding sites at the replication fork. Although the C-terminal region of SUUR is necessary to induce underreplication (Kolesnikova, 2005), the C-terminal portion of SUUR remains associated with heterochromatin in the SUURΔSNF construct, but this protein is not sufficient to induce underreplication. It is suggested that at physiological levels, the affinity of SUUR for replication forks is substantially diminished in the absence of the SNF2 domain. This work raises questions about the biological significance of SUUR binding to heterochromatin, since without the SNF2 domain SUUR is still constitutively bound to heterochromatin, yet unable to induce underreplication. Additionally, SUUR dynamically associates with heterochromatin in mitotic cells although heterochromatin is fully replicated (Munden, 2018).

While trying to uncover the molecular mechanism through which SUUR is able to inhibit replication fork progression, this study has uncovered an interaction between SUUR and Rif1. Through subsequent analysis, it was demonstrated that Rif1 has a direct role in copy number control and that Rif1 acts downstream of SUUR in the underreplication process. Although underreplication is largely dependent on SUUR, there are several sites that display a modest degree of underreplication in the absence of SUUR. In a Rif1 mutant, however, these sites are fully replicated and there is no longer any detectable levels of underreplication within any regions of the genome. It is possible that Rif1 is capable of promoting underreplication through a mechanism independent of SUUR. Therefore, it is concluded that Rif1 is a critical factor in driving underreplication (Munden, 2018).

Further emphasizing the critical role Rif1 plays in copy number control, this study has shown that Rif1 acts downstream of SUUR in promoting underreplication. SUUR is still able to associate with chromatin in the absence of Rif1 but is unable to promote underreplication. Underreplicated regions of the genome, including heterochromatin, tend to be late replicating, raising the possibility that changes in replication timing in a Rif1 mutant suppresses underreplication. Rif1 mutant endocycling cells of Drosophila display a cytological pattern of late replication, where heterochromatin is discretely replicated. While Rif1 controls replication timing in Drosophila and is necessary for the onset of late replication at the mid-blastula transition (Seller, 2018), it is argued that the changes in copy number associated with loss of Rif1 function are not solely due to a loss of late replication. This is supported by the clear pattern of late replication of heterochromatin in Rif1 mutant endocycling cells, although heterochromatin appears to be fully replicated in these cells. Previous work in mammalian polyploid cells has shown that underreplication is dependent on Rif1, which was attributed to changes in replication timing (Hannibal, 2016). It is important to note that Rif1-dependent changes in replication timing were not measured in this system and that many genomic regions transition from early to late replication in a Rif1 mutant (Foti, 2016). This work raises the possibility that Rif1 has a direct role in mammalian underreplication through a mechanism similar to that of Drosophila and may not simply be due to indirect changes in replication timing. Future work will be necessary to define the role of mammalian Rif1 in underreplication (Munden, 2018).

This analysis of amplification loci demonstrates that Rif1 controls replication fork progression independently of initiation control, thus demonstrating that Rif1 has a specific effect on replication fork progression. Therefore, this study has uncovered a new role for Rif1 in DNA metabolism as a regulator of replication fork progression and copy number control. Rif1 has been identified as part of the replisome in human cells by nascent chromatin capture, a technique that identifies proteins associated with newly synthesized chromatin. Multiple studies have assessed whether loss of Rif1 function affects replication fork progression in yeast, mouse and human cells, but have come to different conclusions. DNA fiber assays have been used to measure fork progression in these studies and nearly all have shown that Rif1 mutants have a slight increase in replication fork progression, although not always statistically significant. There could be several reasons for these differing results; Rif1 may control replication fork progression in specific genomic regions that may be underrepresented in some assays, Rif1 function could vary among different cell types, or sample sizes may have been too small to reach significance. These observations, taken together with these previous studies, leave open the possibility that Rif1-mediated control of replication fork progression could be an evolutionarily conserved function of Rif1. It is not suggested that Rif1 is constitutively associated with replication forks in all cell types. Rather, Rif1 could be recruited to replication forks at a specific time in S phase, or in specific developmental contexts, to modulate the progression of replication forks and provide an additional layer of regulation of the DNA replication program (Munden, 2018).

How could SUUR and Rif1 function in concert to inhibit replication fork progression? This study has shown that Rif1 retention at replication forks is dependent on SUUR. Additionally, underreplication depends on Rif1's PP1-binding motif, raising the possibility that a Rif1/PP1 complex is necessary to inhibit replication fork progression. Rif1/PP1 dephosphorylates DDK-activated helicases to control replication initiation. Recently, however, DDK-phosphorylated MCM subunits have been shown to be necessary to maintain DNA-unwinding enzyme Cdc45~MCM~GINS (CMG) association and stability of the helicase (Alver, 2017). This result suggests that continued phosphorylation of the helicase is necessary for replication fork progression (Alver, 2017). It is proposed that SUUR recruits Rif1/PP1 to replication forks where it is able to dephosphorylate MCM subunits, ultimately inhibiting replication fork progression. Although this mechanism needs to be tested biochemically, it provides a framework to address the underlying molecular mechanism responsible for controlling DNA copy number and could provide new insight into the mechanism(s) Rif1 employs to regulate replication timing. (Munden, 2018).

list of proteins involved in DNA replication


Andreyeva, E. N., Bernardo, T. J., Kolesnikova, T. D., Lu, X., Yarinich, L. A., Bartholdy, B. A., Guo, X., Posukh, O. V., Healton, S., Willcockson, M. A., Pindyurin, A. V., Zhimulev, I. F., Skoultchi, A. I. and Fyodorov, D. V. (2017). Regulatory functions and chromatin loading dynamics of linker histone H1 during endoreplication in Drosophila. Genes Dev 31(6): 603-616. PubMed ID: 28404631

Armstrong, R. L., Penke, T. J. R., Strahl, B. D., Matera, A. G., McKay, D. J., MacAlpine, D. M. and Duronio, R. J. (2018). Chromatin conformation and transcriptional activity are permissive regulators of DNA replication initiation in Drosophila. Genome Res. PubMed ID: 30279224

Brush, G. S. and Kelly, T. J. (1996). Mechanisms for replicating DNA. In "DNA replication in eukaryotic cells". Ed. M. L. DePamphilis. pp 1-43 Cold Spring Harbor Laboratory Press, Plainview, NY.

Brustel, J., Kirstein, N., Izard, F., Grimaud, C., Prorok, P., Cayrou, C., Schotta, G., Abdelsamie, A. F., Dejardin, J., Mechali, M., Baldacci, G., Sardet, C., Cadoret, J. C., Schepers, A. and Julien, E. (2017). Histone H4K20 tri-methylation at late-firing origins ensures timely heterochromatin replication. EMBO J 36(18): 2726-2741. PubMed ID: 28778956

Chong, J. P. J., Thömmes, P and Blow, J. J. (1996). The role of MCM/P1 proteins in licensing of DNA replication. Trends in Biochem. Sci. 21: 102-106. Medline abstract: 8882583

Hiratani, I., et al. (2008). Global reorganization of replication domains during embryonic stem cell differentiation. PLoS Biol. 6: e245. PubMed Citation: 18842067

Hou, C., Li, L., Qin, Z. S. and Corces, V. G. (2012). Gene density, transcription, and insulators contribute to the partition of the Drosophila genome into physical domains. Mol Cell 48(3): 471-484. PubMed ID: 23041285

Hua, B. L., Bell, G. W., Kashevsky, H., Von Stetina, J. R. and Orr-Weaver, T. L. (2018). Dynamic changes in ORC localization and replication fork progression during tissue differentiation. BMC Genomics 19(1): 623. PubMed ID: 30134926

Kolesnikova, T. D., Goncharov, F. P. and Zhimulev, I. F. (2018). Similarity in replication timing between polytene and diploid cells is associated with the organization of the Drosophila genome. PLoS One 13(4): e0195207. PubMed ID: 29659604

Lucas, I., et al. (2007). High-throughput mapping of origins of replication in human cells. EMBO Rep. 8: 770-777. PubMed Citation: 17668008

Munden, A., Rong, Z., Sun, A., Gangula, R., Mallal, S. and Nordman, J. T. (2018). Rif1 inhibits replication fork progression and controls DNA copy number in Drosophila. Elife 7. PubMed ID: 30277458

Nordman, J. T., Kozhevnikova, E. N., Verrijzer, C. P., Pindyurin, A. V., Andreyeva, E. N., Shloma, V. V., Zhimulev, I. F. and Orr-Weaver, T. L. (2014). DNA copy-number control through inhibition of replication fork progression. Cell Rep 9: 841-849. PubMed ID: 25437540

Schwaiger, M., Stadler, M. B., Bell, O., Kohler, H., Oakeley, E. J. and Schubeler, D. (2009). Chromatin state marks cell-type- and gender-specific replication of the Drosophila genome. Genes Dev 23(5): 589-601. PubMed ID: 19270159

Torres-Rosell, J., et al. (2007). Anaphase onset before complete DNA replication with intact checkpoint responses. Science 315: 1411-1415. PubMed Citation: 17347440

Yarosh, W. and Spradling, A. C. (2014). Incomplete replication generates somatic DNA alterations within Drosophila polytene salivary gland cells. Genes Dev 28(16): 1840-1855. PubMed ID: 25128500

Zhimulev, I. F., Zykova, T. Y., Goncharov, F. P., Khoroshko, V. A., Demakova, O. V., Semeshin, V. F., Pokholkova, G. V., Boldyreva, L. V., Demidova, D. S., Babenko, V. N., Demakov, S. A. and Belyaeva, E. S. (2014). Genetic organization of interphase chromosome bands and interbands in Drosophila melanogaster. PLoS One 9: e101631. PubMed ID: 25072930

date revised: 25 October 2017

Zygotically transcribed genes

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